185-199 - Revista Iberoamericana de Micología

Transcription

Rev Iberoam Micol. 2012;29(4):185–199
Revista Iberoamericana
de Micología
www.elsevier.es/reviberoammicol
Review
The taxonomy and phylogenetics of the human and animal pathogen
Rhinosporidium seeberi: A critical review
Raquel Vilela a,c,d , Leonel Mendoza a,b,∗
a
Biomedical Laboratory Diagnostics, Michigan State University, East Lansing, MI 48424-1031, USA
Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI 48424-1031, USA
Faculty of Pharmacy, Federal University of Minas Gerais, Belo Horizonte, Brazil
d
Institute Superior of Medicine (ISMD), Minas Gerais, Belo Horizonte, Brazil
b
c
a r t i c l e
i n f o
Article history:
Received 15 February 2012
Accepted 26 March 2012
Available online 12 April 2012
Keywords:
Rhinosporidium seeberi
Rhinosporidiosis
Mesomycetozoa
Plastids
Phylogenetics
a b s t r a c t
Rhinosporidum seeberi is the etiologic agent of rhinosporidiosis, a disease of mucous membranes and
infrequent of the skin and other tissues of humans and animals. Because it resists culture, for more than
100 years true taxonomic identity of R. seeberi has been controversial. Three hypotheses in a long list of
related views have been recently introduced: 1) a prokaryote cyanobacterium in the genus Microcystis
is the etiologic agent of rhinosporidiosis, 2) R. seeberi is a eukaryote pathogen in the Mesomycetozoa
and 3) R. seeberi is a fungus. The reviewed literature on the electron microscopic, the histopathological
and more recently the data from several molecular studies strongly support the view that R. seeberi is
a eukaryote pathogen, but not a fungus. The suggested morphological resemblance of R. seeberi with
the genera Microcystis (bacteria), Synchytrium and Colletotrichum (fungi) by different teams is merely
hypothetical and lacked the scientific rigor needed to validate the proposed systems. A fundamental
aspect against the prokaryote theory is the presence of nuclei reported by numerous authors and updated
in this review. Moreover, Microcystis’s and Synchytrium’s ultra-structural and key cell cycle traits cannot
be found in R. seeberi parasitic phase. The PCR amplification of a cyanobacteria 16S rDNA sequence from
cases of rhinosporidiosis, while intriguing, will be viewed here as an anomaly due to contamination with
environmental Microcystis or perhaps as an endosymbiotic acquisition of plastids from cyanobacteria
ancestors. Thus, even if R. seeberi possesses prokaryote DNA, this does not prove that R. seeberi is a
cyanobacterium. The placement of R. seeberi within the fungi is scientifically untenable. The isolation and
the DNA analysis performed in a fungal strain, and the lack of appropriate controls are the main problems
of this claim. Further studies are needed to validate R. seeberi’s acquisition of prokaryote plastids and other
issues that still need careful scrutiny.
© 2012 Revista Iberoamericana de Micología. Published by Elsevier España, S.L. All rights reserved.
Taxonomía y filogenética de Rhinosporidium seeberi, un patógeno para el ser
humano y los animales: revisión crítica
r e s u m e n
Palabras clave:
Rinosporidiosis
Mesomycetozoa
Plástidos
Filogenética
Rhinosporidum seeberi es el agente etiológico de la rinosporidiosis, una enfermedad de las membranas
mucosas y, con menos frecuencia, de la piel y otros tejidos. Debido a que se resiste a crecer en los
medios de cultivo desde hace más de 100 años, la identidad taxonómica de R. seeberi ha sido motivo
de controversia. Tres nuevas hipótesis en una larga lista de puntos de vista similares han sido introducidas: 1) la cianobacteria Microcystis es el agente etiológico de la rinosporidiosis, 2) R. seeberi es un
patógeno eucariota en los Mesomycetozoa, y 3) R. seeberi es un hongo. La literatura revisada sobre
los estudios realizados con microscopia electrónica, los datos histopatológico y, más recientemente,
los datos de varios estudios moleculares, apoyan fuertemente la idea de que R. seeberi es un patógeno
eucariota, pero no un hongo. La semejanza morfológica propuesta por algunos de que R. seeberi es
similar a los miembros de los géneros Microcystis (bacteria), Synchytrium y Colletotrichum (hongos)
es meramente hipotética y no tiene el rigor científico necesario para validar el sistema propuesto.
Un aspecto fundamental en contra de la teoría procariota es la presencia de núcleos descrita por
numerosos autores y que actualizamos en esta revisión. Además, las características ultra-estructurales
∗ Corresponding author.
E-mail address: mendoza9@msu.edu (L. Mendoza).
1130-1406/$ – see front matter © 2012 Revista Iberoamericana de Micología. Published by Elsevier España, S.L. All rights reserved.
http://dx.doi.org/10.1016/j.riam.2012.03.012
186
R. Vilela, L. Mendoza / Rev Iberoam Micol. 2012;29(4):185–199
de los géneros Microcystis y Synchytrium y de sus ciclos celulares no han sido encontradas en la fase parasitaria de R. seeberi. La amplificación por PCR de una secuencia del rADN 16S típica de las cianobacterias
en muestras de casos de rinosporidiosis, aunque interesante, será considerada en esta revisión como una
anomalía debido a la contaminación con el medio ambiente (Microcystis) o tal vez como una adquisición
endosimbiótica de plastidios a partir de cianobacterias ancestrales. Así pues, aunque R. seeberi podría
poseer ADN procariota, esto no demuestra necesariamente que R. seeberi sea una cianobacteria. La clasificación de R. seeberi dentro de los hongos es insostenible. El aislamiento de un hongo, los análisis de ADN
realizados, y la ausencia de controles apropiados son los problemas más importantes de esta teoría. Más
estudios serán necesarios para validar la adquisición de plastidios procariotas en R. seeberi, y otros temas
que requieren un cuidadoso escrutinio.
© 2012 Revista Iberoamericana de Micología. Publicado por Elsevier España, S.L. Todos los derechos
reservados.
Rhinosporidium seeberi is the causative agent of rhinosporidiosis
in humans and animals.14,59,100 The disease affects mucous membranes and rarely the skin and/or the internal tissues of its infected
hosts.14,60 The first report of rhinosporidiosis was published by
Seeber88 and occurred in a 19-year-old Argentinean patient with
breathing difficulties caused by a polyp that obstructed his nasal
passages. In his thesis Seeber88 noted also that in 1892 Malbram,
in Buenos Aires, Argentina, was the first to diagnose the disease in
humans, but he did not publish his finding. Although the disease has
been frequently diagnosed in the Americas and other areas of world,
rhinosporidiosis is more prevalent in India and Sri Lanka than in
any other geographic locations.14,100 The original published cases
in Argentina, and many others recorded thereafter, established that
R. seeberi develops in the infected hosts 10 to >450 ␮m in diameter spherical structures (sporangia), some containing hundreds of
endoconidia (sporoblast, trophocyte), as well as immature forms
of different sizes (5–80 ␮m) without endoconidia. Despite the large
numbers of spherical structures in the infected tissues, it was soon
apparent that R. seeberi resists culture and that a rhinosporidiosis
infection cannot be induced in experimental animals.14,15 These
unique features steamed a great controversy that still continues to
the present days.9,35,42
As has been the case with a number of uncultivated pathogens,
such as Lacazia loboi61,102 and Mycobacterium leprae,66,78,81 numerous investigators have claimed that they had successfully produced
R. seeberi in culture.27,58,65 In general most of the isolated strains
were later found to be common contaminants.34,66,103 A good
example of this trend is the most recent report claiming that
R. seeberi is not a mesomycetozoa or a bacterium, but a fungus
(97). This new proposal was based on fungal strains recovered
from cases of rhinosporidiosis and from a frozen clinical sample
collected ten years ago from a patient with rhinosporidiosis in
India (97, 98). Thus, strains obtained from well known uncultivated pathogens have always been suspect and their identities are
subjected to endless controversies. However, with the advent of
molecular tools most of these debates came to an end, although
not for R. seeberi.9,25,42,69,103
Currently there are three main hypotheses on the taxonomic
and phylogenetic positions of R. seeberi in the tree of life; the
first argues that R. seeberi is a prokaryote cyanobacterium in the
genus Microcystis,6,9 the second one claims that R. seeberi is a
eukaryote microbe closely related to spherical fish pathogens in
the Mesomycetozoa,35,42 and the most recent hypothesis claims
that R. seeberi is a fungus. These conflicting views on the taxonomy and phylogeny of R. seeberi will be critically addressed in
this review. To put into perspective these hypotheses, we will first
review results from traditional studies of R. seeberi to define its
main characteristics and its taxonomy by focusing on the histological and electron microscopy (EM) data that have been accumulated
over the last 110 years. We will also provide a historical description
of the current R. seeberi theories to finally critically address these
positions.
Traditional nomenclature, systematic and current views
on R. seeberi
The taxonomic classification of R. seeberi has been always contentious. Based on its parasitic spherical morphological features
with the development of endoconidia, Seeber88 believed that it
was a protist closely related to a Coccidium (Coccidium seeberia, Wernecki).16,20 Sebeer’s former adviser Wernicke20 thought it
could also be a fungus similar to Coccidium immitis, and Minchin and
Fantham72 grouped the pathogen with the Haplosporidia. Battie19
classified it with the Neurosporidia, and Ashworth16 treated it
as a fungus in the zygomycetes or possibly a chytridiomycete.
Based on ultrastructural characteristics, Vanbreuseghem101 suggested that R. seeberi may have a relationship with the algae,
whereas Laveran and Petit63 were the first to note the resemblance of Ichthyosporidum (currently Icthyophonus a pathogen of
trout) with R. seeberi. Later Acevedo2 and Carini22 mentioned that
R. seeberi also shared morphological attributes with Dermosporidium hylarum (Amphibiocystidium ranae).82 Interestingly, these two
pathogens of fishes and frogs are currently classified with members
of the Dermocystida in the Mesomycetozoa (see below).3,68
When Seeber88 recorded the first case of rhinosporidiosis, he
carefully described R. seeberi’s morphological features in infected
tissues, but did not name the organism. Instead the resemblance of this pathogen with C. immitis, the etiologic agent of
coccidioidomycosis,105 was mentioned. It is believed that Seeber’s
professor Robert Wernicke named the organism C. seeberia, as
appeared in a small note on the “Tratado de Parasitología Animal” in 1903.20 Apparently, Wernicke suggested also the name
“Coccidium seeberi” in an ephemeral publication in 1900 at the
University of Buenos Aires, Argentina under the name “Programa
de Zoología Médica”.16,32 Even though the 1900 original edition
was no longer available, Ashworth16 acquired a copy of the document dated 1907 and stated that the species C. seeberi was validly
introduced. Unaware of these proposals, Minchin and Fantham72
studied histological samples of an Indian rhinosporidiosis case,
described earlier by O’Kinealy,77 and introduced the binomial Rhinosporidium kinealyi. Soon, other cases of rhinosporidiosis were
also attributed to R. kinealyi, including the first case of the disease in
the United Sates.19,107 In response to Minchin and Fantham’s72 proposal, Seeber89 adopted the genus Rhinosporidium, but argues for
the species priority of the original name.20 He suggested that the
combination R. seeberi should be accepted. In 1913 Zschokke110
introduced yet another species Rhinosporidium equi after studying histological sections obtained from a horse with a nasal polyp
in South Africa. However, in 1923 Ashworth16 revised all proposals and concluded that R. seeberi has priority over new proposed
species. Thus, the species Rhinosporidium ayyari,11 Rhinosporidium hylarum,2,22 and Rhinosporidium amazonicum1 introduced later
became synonyms of R. seeberi.
Bizarre hypotheses claiming that the spherical structures
found in rhinosporidiosis were self-assembled material made of a
mixture of plant and human material or spheres of cellular waste
due to the ingestion of tapioca, have been suggested.7,10,17 However, these extreme views were soon abandoned. Subsequently
Alhuwalia et al.9 proposed a theory linking R. seeberi with a
prokaryote cyanobacterium in the genus Microcystis, a view that
challenged traditional studies placing R. seeberi among the eukaryote microbes.14 Concurrently, Herr et al.,42 and Fredricks et al.,35
based on morphological and the PCR amplification of the 18S small
subunit ribosomal DNA sequences, concluded that this pathogen
was an eukaryote microbe. Herr et al.,42 introduced the name
Mesomycetozoa (between animals and fungi) to place R. seeberi
with microbes displaying spherical phenotypes with endoconidia,
such as Dermocystidium and Sphaerothecum destruens (the rosette
agent). Based on molecular phylogenetic analysis methods and the
available morphological data, the possibility of several host specific strains in the genus Rhinosporidium was also suggested.91
Thankamani99 and Thankamani and Lipin-Dev97 introduced the
most recent hypothesis on the taxonomic placement of this unique
pathogen. They claimed the isolation of a fungus from cases of rhinosporidiosis and thus, they believe this fungal strain must be the
etiologic agent of rhinosporidiosis.
Historical studies on the parasitic life cycle of R. seeberi
Seeber88 was the first to address R. seeberi’s complex life
cycle in the tissue of its infected hosts. His observations were
later confirmed by numerous investigators studying the morphological characteristics of R. seeberi in humans and animals with
rhinosporidiosis.14 After Seeber’s thesis,88 several comprehensive
studies on the morphological and life cycle features of this unique
pathogen were published by Acevedo,2 Ahsworth,16 Grover,39
Kurunaratne,59 Melo,67 and Thianprasit and Thagerngpol.100
Because R. seeberi resists culture, its true ecological niche is still
unknown.16,26,59 Based on the proximity of the affected patients to
wet environments, most investigators agree with the concept that
R. seeberi must be located in aquatic niches. Although the occurrence of rhinosporidiosis in dry areas of the Middle East seems to
contradict this view, it is quite possible that R. seeberi had evolved
also to develop resistant structures in dry environments.14 Despite
the finding of rhinosporidiosis cases in arid areas, it is widely
accepted that the infecting units (possibly non-flagellate spores)
are more likely located in aquatic environments and may contact
affected hosts through traumatic lesions, leading to the implantation of the pathogen in the host’s tissues (see below).26 Once
in the host, the infecting unit of unknown origin increases in size
(10 to >450 ␮m in diameter) and through cytoplasmic cleavage produces hundreds of endoconidia within the spherical cells, which
subsequently are released through a single pore and reinitiate a
life cycle.47,54
According to numerous studies using stained tissue sections and
electron microscopy (EM),2,18,39,45,48,53,75,95,96 R. seeberi’s life cycle
starts with the release of mature endoconidia through a small pore
in the cell wall of the mature sporangium (Fig. 1, this review). A film
depicting R. seeberi endoconidia release through a pore upon activation with water is available at: http://www.bld.msu.edu/Rhino.
The diameter of an endoconidia ranges from 4 to 10 ␮m. Numerous EM studies have also revealed that the endoconidia possess a
well defined thin bilaminated cell wall (1–3 ␮m) containing several small vesicles termed electron dense bodies (EDBs), a nucleus
with a distinctive nucleolus, and a granular mucilaginous capsule,
as those also shown in Fig. 2A, this review.2,16,47,54,59,75,82,87,95,101
Some investigators have further postulated that the EDBs within
the endoconidium are the true generative infective units of
R. seeberi and even have proposed the term “spore-morula”
for the way these structures give origin to endosporulated
187
Fig. 1. Electron microscopy tissue section showing a Rhinosporidium seeberi
mature sporangium containing hundreds of endoconidia, one already outside
the sporangium and the others in the process of being expelled through a
pore (arrow heads) (a video depicting the endoconidia release is available at:
http://www.bld.msu.edu/Rhino). Note the presence of a thin cell wall and the formation of three prominent inner layers (a clear space between the mature endoconidia
and the cell wall) primarily located near the pore. At this magnification this structure
appears as a single inner layer, but it comprises three well defined inner layers.40 The
presence of fully developed endoconidia is observed at the center and toward the
pore, whereas immature small usually oval endoconidia are found at the opposite
site (white heads), a distinctive feature of mature sporangia. Three immature sporangia are also noted in the lower section of the mature sporangium (white arrow
heads) (Bar = 100 (␮m).
sporangia.12,19,60 However, this theory has few followers and so far
has not been fully accepted. Although some authors have reported
the presence of nuclei and mitochondria in the endoconidia, these
two organelles are extremely difficult to visualize either before or
after the release of the endoconidia (Fig. 2A, this review). This is
probably due to problems during the tissue fixative process prior
to staining protocols.16,40,45,59,87,95,101,106
It has been postulated that after the release, the endoconidia
increase in size (∼10–70 ␮m) and by absorption loses all vesicles and EDBs to become a juvenile sporangium (JS) (Fig. 2B, this
review).16,47,54,59,87 Based on EM studies, this stage is characterized by a prominent cell wall, a granular cytoplasm, the presence
of a single central nucleus with a noticeable nucleolus enclosed
by nuclear membrane, and some mitochondria (Figs. 2B and 3A,
this review).2,16,46,54,59,75,87,101 The presence of endoplasmic reticulum, lipidic globules, and vacuolate structures has also been
described.40,45,54,87 The juvenile sporangium increases in size and
becomes an intermediate sporangium (IS) reaching ∼70–150 ␮m
in diameter (Fig. 3B, this review). Several investigators, including Acevedo,2 Ahsworth,16 Herr et al.,40 Kurunaratne,59 and others
have reported at this stage the presence of a thick cell wall containing chitin, granular cytoplasm with numerous nuclei dispersed
within the cytoplasm, numerous flat cristae mitochondria, the presence of an early pore, and one or more electron dense concentric
rings termed “laminated bodies”. The latter structures have been
considered by some as artifacts due to the tissue fixative process,67
or as structures made of chromatin by others.42,87 The presence
of immature endoconidia at this stage is a rare finding. By nuclear
cleavage the last nuclear division takes place. This event is followed
by the development of a thin cell wall around each nucleus, giving
rise to the formation of immature endoconidia, and thus the whole
structure becomes an early mature sporangium.17,18,45,47 Most of
the morphological features of mature sporangia such as the presence of inner layers and a visible pore are missing at this early
stage.
188
Fig. 2. This electron micrograph shows several Rhinosporidium seeberi’s purified
endoconidia, one of them containing a nucleus with a prominent nucleolus (arrow)
(Panel A). The endoconidia are characterized by a thin cell wall, numerous electron
dense bodies (EDBs, dark vesicles), and the presence of a granular mucilaginous
capsule (Bar = 5 (␮m). Panel B shows three early juvenile sporangia (JS) within the
host infected tissues, before the first nuclear division (Bar = 15 (␮m). The presence
of nuclei with prominent nucleolus is observed in two of the JS (arrow heads), which
contrast with the nucleus displayed by the host’s inflammatory cells (arrow). The
JS on the left lacks nucleus because it was sectioned in a segment of the cell where
the nucleus was not near. The presence of prominent cell wall and several vesicles
typical of this stage is also noted.
Mature sporangia can reach 450 ␮m or more in diameter (Fig. 1,
this review). At this stage the sporangium is readily identified by
the presence of hundreds of mature and immature endoconidia,
by a thin cell wall containing at least three electron lucid and
antigenic inner layers, the presence of an exit pore, and by its
enormous size. The site of the exit pore (∼10–20 ␮m in diameter) is believed to be genetically determined71 and is always facing
the mature spherical endoconidia containing numerous vesicles,
whereas the opposite site contains mostly small oval immature
endoconidia (2–5 ␮m in diameter) with few vesicles, and thus it
has been referred by some as the “germinative zone” (Fig. 1, this
review).87 Mendoza et al.,71 and others18,65,87 reported the formation of new endoconidia directly from this particular area. It has also
been shown by Ashworth16 and Mendoza et al.71 that watery substances facilitate the release of mature endoconidia. These authors
further proposed that upon contact with watery substances lytic
enzymes are accumulated at the pore site, a feature that could facilitate the release of the endoconidia due to an increase in the osmotic
Fig. 3. Panel A shows a histopathological section of Rhinosporidium seeberi stained
with H&E depicting an early juvenile sporangium (JS) similar to the one depicted
in Fig. 2 Panel B (Bar = 8 (␮m). The presence of a well delimited cell wall, a granular cytoplasm with a typical reddish nucleus and a prominent nucleolus is typical
of this stage. The JS are devoid of vesicles and EDBs, structures typically found in
the endoconidia within or outside mature sporangia. The nucleus’ reddish color
sharply contrasts with the hosts’ bluish nuclei surrounding the JS. Panel B depicts an
intermediate sporangium with a thicker cell wall and several reddish nuclei, before
nuclear division, identical to the one displayed by the JS in Panel A (Bar = 20 (␮m).
Note the contrast between R. seeberi’s reddish nuclei and the host nuclei. This stage
is characterized by multiple synchronized nuclear divisions.
pressure in the mature sporangia. This mechanism first proposed
by Ashworth,16 was later adopted by Mendoza et al.,71 to explain
the methodical release of the endoconidia from mature sporangia. A
summary of R. seeberi life cycle, based on the accumulated thoughts
and observations, is presented in Fig. 4 of this review.
Relevant to R. seeberi developmental features within the spherical cells are the long forgotten depictions of mitotically dividing
nuclei and the presence of chromosomes, as reported by the early
investigators.2,16,59 The presence of nuclei in the parasitic stages
of R. seeberi was first recorded by Seeber,84 who on page 35 of his
thesis depicted a nucleus within the “sporoblast” (endoconidium),
a finding corroborated later by others.2,16,19,42,46,59,82,96,100 The first
description of mitotic figures within immature sporangia, however,
was that of Ashworth,16 who provided (Plate 1, Figs. 3–17) illustrations of the mitotic figures he observed of prophase and telophase
nuclei during the first nuclear division and the presence of at least
four chromosomes. The first comprehensive photographic description on the histological events of nuclear division within the JS and
IS was presented in the thesis of Acevedo2 (pages 66–73), where
he depicts several single nuclei and the first nuclear division within
the endoconidia (Figs. 41, 42, 44). In addition, he also shows in
Figs. 45–49 and 56–61 the presence of multiple nuclei and nuclear
divisions within immature sporangia, similar to that depicted in
Fig. 4. The figure depicts the parasitic life cycle of Rhinosporidium seeberi based on
data published in the last 110 years. It starts with the release of the endoconidia
through a pore from mature sporangia (a). The released endoconidia (b) increase in
size (10–70 (␮m) loosing the morphological features typical of endoconidia such as
vesicles and EDBs, becoming juvenile sporangia (JS) (c). This is the most commonly
found phenotype in histopathological preparations. The JS are characterized by the
presence of granular cytoplasm, reddish (in H&E) nuclei (could be a single nucleus
or more than two nuclei) with a prominent nucleolus similar to those displayed in
Fig. 3, and a thick cell wall. The JS increases in size (∼70–150 (␮m) and becomes
intermediate sporangia (IS) (d). This is a rare encountered phenotype. The stage
is characterized by synchronized nuclear divisions developing several hundreds of
new nuclei without the formation of a cell wall within the sporangia. The sporangia
at this stage possess a thick cell wall. The presence of rudimentary pore can be found
at this stage, but it is a very rare event, thus difficult to find in histological sections. As
the nuclei multiply, the IS continue increasing in size (≥150 (␮m) becoming an early
mature sporangia (e). At this stage the last synchronized nuclear division takes place,
after which a thin cell wall becomes apparent around each nucleus. Eventually, the
sporangia reach maturity which is followed by the formation of a pore allowing the
release of the endoconidia and the cell cycle is repeated (a).
Fig. 3B of this review. Acevedo’s observations2 were validated six
years later by Kurunaratne.59
In the first book ever published on “Rhinosporidiosis in Man” (an
update of a previous edition), Karunaratne59 thoroughly reviewed
all aspects related to R. seeberi including its parasitic cell cycle.
Karunaratne59 showed photographs (Figs. 1–14) of the first nuclear
division and the presence of four chromosomes (as predicted
by Ashworth16 ) within the early JS and numerous nuclei within
IS during prophase, metaphase and anaphase. With few exceptions, subsequent studies did not explore this fascinating feature
of R. seeberi and mostly concentrated on other aspects of its life
cycle.17,18,42,46,47,54,65,73,75,83,87,96,100,101
R. seeberi’s eukaryote and prokaryote theories
R. seeberi is a prokaryote cyanobacterium within the genus Microcystis. First Alhuwalia et al.,9 and then Alhuwalia6 reported that
they have evidence indicating that R. seeberi is a cyanobacterium
in the genus Microcystis. This hypothesis was unveiled in India
based on ecological studies conducted in ponds and rivers where
patients with rhinosporidiosis often bath. Environmental discoid
and oval structures that, according to these authors, resembled
the parasitic stage of R. seeberi in the infected tissues of patients
with rhinosporidiosis were found. They stated that the cocci
(nanocytes) developed by the genus Microcystis in these aquatic
environments possessed striking morphological similarities with
R. seeberi’s endoconidia and thus, these structures may be the same
189
elements observed in cases of rhinosporidiosis. They proposed that
the nanocytes might be the infecting units of the diseases rhinosporidiosis. They also suggested that the amorphous and oval
colonies filled with numerous internal cocci of Microcystis found in
the ponds, were the same as mature sporangia with endoconidia
observed in the infected tissues of patients with rhinosporidiosis.
However, these observational data were not further investigated to
validate the author’s hypothesis.
The initial investigations putting forth the Microcystis hypothesis were followed by a study reporting its isolation using BG11
medium (to isolate cyanobacteria) and a slide culture technique
with different concentrations of glycerol.9 According to the authors
of this study described what they contend is a reliable method
to culture R. seeberi. They also reported the isolation of Microcystis from clinical cases of rhinosporidiosis, and from water ponds,
the detection of spherical structures containing nanocytes in BG11
medium and the formation of some filaments identified as nonfungal in origin in the slide cultures. Based on their findings, the
authors concluded they had recovered from both, natural sources
and clinical samples of patients with rhinosporidiosis, a cyanobacterium of the genus Microcystis and thus that this microbe was the
legitimate etiologic agent of rhinosporidiosis.
Dhaulakhandi et al.,30 using molecular methodologies amplified
a 16S rRNA sequence (accession number AJ440719) from purified
single cells of R. seeberi obtained from patients with rhinosporidiosis and from environmental samples. The rRNA sequence contained
the typical AATTTTCCG signature for cyanobateria and two other
palindromic repeats present in Microcystis species, including Microcystis aeruginosa. Since the round bodies (RBs) of R. seeberi resemble
Microcystis phenotypes and the 16S rRNA sequence has strong DNA
identity with other cyanobacteria, they concluded that the etiologic
agent of rhinosporidiosis is a cyanobacterium and thus, R. seeberi
should be reclassified.70 More recently, Alhuwalia4 conducted new
phylogenetic studies with the sequence obtained from their previous study and found that the 16S rRNA R. seeberi sequence has 98%
identity with flowering plants and only 79–86% identity with Microcystis and with other cyanobacteria in the chroccocales. Despite
this result, they argued that the morphological features displayed
by R. seeberi in patients with the disease have more characteristics in common with the cyanobacteria than with the eukaryotes
microbes in the Mesomycetozoa.4
In a letter to the editor, Alhuwalia5 defended her position arguing that the mistake leading Herr et al.42 to classify R. seeberi as
a eukaryote microbe in the Mesomycetozoa, was the use of nonpurified sporangia and endoconidia from the infected tissues of
humans with rhinosporidiosis. She concluded that the amplified
DNA sequence of Herr et al.,42 was probably of human origin, a
claim properly refuted by the authors.70 She also stated that the
structures in the EM figures identified by these authors as nuclei
were “not nuclei but nanocytes of Microcystis encompassing naked
prokaryotic DNA”. She further contended that the structures named
as mitochondria by Herr et al.,42 were of human origin and not
from R. seeberi sporangia. She based this conclusion on numerous
EM studies that did not find mitochondria in young and mature
sporangia.
R. seeberi is a eukaryote microbe within the Mesomycetozoa. In
1999 Herr et al.,42 provided taxonomic and molecular data indicating that R. seeberi is an eukaryote microbe within a new cluster
of aquatic pathogens, first described by Ragan et al.84 as the DRIP
group (after Dermocystidium, Rosetta Agent = currently S. destruens,
Ichthyophonus, and Psorospermium), and renamed by Herr et al.42
as the Mesomycetozoa3 (Coanozoa of Cavalier-Smith23 ). This finding was welcome by the scientific community in part, because
it corroborated 100 years of studies linking this pathogen to the
eukaryotes.16–18,45–47,53,54,65,75,83,87,88,100,101 To reach their conclusion, Herr et al.,42 collected tissue samples from two Sri Lankan
190
men with rhinosporidiosis and conducted EM and molecular studies. The EM studies of the infected tissues showed mature and
immature sporangia of different sizes, some nuclei containing a
prominent nucleolus and with numerous flat mitochondrial cristae
detected in intermediate sporangia. Although this position was
rejected as inaccurate by Alhuwalia5 (see above), in a response
to her critique Mendoza et al.,70 provided additional EM photos
substantiating their findings.
In the above study the tissue samples were processed in two
ways: a) the sporangia were mechanically dissected from the
infected tissues washed with sterile distilled water and then disrupted in the presence of glass beads, and b) the tissue samples
were placed in a mortar and ground while frozen in the presence
of liquid nitrogen with a pestle. The genomic DNA extracted from
these samples was then subjected to PCR amplification using the
universal primers NS1 and NS8.42 R. seeberi 18S SSU rDNA 1790 bp
PCR DNA fragment was amplified (accession numbers: AF118851).
Phylogenetic analysis of the 18S SSU rDNA sequence and 23 other
DNA sequences of different microbes showed R. seeberi as the sister
taxon to Dermocystidium species and closely related the members
in the Ichthyophonida (see below).
Almost concomitantly, Fredricks et al.35 published an identical study claiming that they had sequenced the 18S SSU rDNA of
R. seeberi from a dog with rhinosporidiosis. These investigators
extracted genomic DNA from the same dog tissue sample used by
Levy et al.65 In addition, they tested formalin fixed tissue samples
of three humans with rhinosporidiosis, and nasal polyps from 12
human patients without the disease as negative controls. In this
new study the design of several primers, including R. seeberi specific primers (Rhino-fw/Rhino-rev) based on DNA unique regions,
were tested and a DNA biotinylated probe for fluorescent in situ
hybridization (FISH) analysis was constructed. The results showed
that the 18S SSU rDNA sequence amplified by Fredricks et al.,35
from the sample from a dog with rhinosporidiosis was identical
to that reported by Herr et al.42 The specific primers designed
to amplify only R. seeberi also failed to produce amplicons using
genomic DNA of humans, fungi, Dermocystidium salmonis, and S.
destruens (rosette agent). Moreover, the R. seeberi specific DNA FISH
probe detected R. seeberi’s DNA present in immature and immature sporangia (blue color), but the negative control probe showed
only auto-fluorescence green (FITC) or red (Texas red) background
according to the used wave-length filter. Of interest was the finding in EM of tubular mitochondrial cristae in R. seeberi’s sporangia,
a finding that contrasted with that of Herr et al.42 Nonetheless,
the authors concluded that their study confirmed that R. seeberi is
not a fungus, but instead it is an unusual aquatic protistan parasite
located near the divergence between animals and fungi, thus validating the findings of Herr et al.42 They further stated that the
specific primers and the FISH DNA probe they developed could
be used to specifically detect this pathogen in putative cases of
rhinosporidiosis.
R. seeberi is a fungus associated to the genera Synchytrium and
Colletotrichum (Glomerella). In 2005 Thankamani99 announced
that she has recovered in culture, from cases of rhinosporidiosis
a fungal organism with “strong resemblance to the developmental
stages of Synchytrium endobioticum, a member of chytridiales that
causes black warts disease in potato”.99 In this preliminary report,
Thankamani99 claimed that the same organism was consistently
isolated from 15 swabs collected in 15 different Indian patients with
rhinosporidiosis. The isolated organism (later labeled as UMH.48)
grew very slow on blood and nutrient agar at room temperature,
but failed to develop at 37 ◦ C. She described the colony recovered at
room temperature as a very small (“pin tip size”) convex, circular,
smooth, semitransparent colony without a diffusible pigment.
When the primary isolate was subculture on Sabouraud dextrose
agar (SDA) the formation of large sporangia with a “narrow
pointing weak area, possibly, developing into a pore. . .” was
noted.99 The author illustrates her findings with several figures
showing the developmental stages of the spherical cells. These
cells grew into spherical bridged cells containing mycelia-like
structures and finally, by multiple divisions, developed into spherical spores. She stated that the formation of mycelia-like structures
was a transient phenomenon leading to spore formation.99 After
repeated subculturing, large sporangia could not be found. The
formation of empty structures (prosorus-like, see below) was
found attached to a homogeneous mass of nuclear material.
Thankamani99 reported also the presence of zoospore-like
structures, but there were no description of their morphological features including the number of flagella and if the cells
were motile. In Fig. 18 the author described “zoospores germinating/flagellated/sexual union of gametes forming zygospore”.99
However, since the flagellum cannot be Gram stained, the figure seems to display only some hyphal elements and a “spore”
with a germ tube. She mentioned also that this strain was used
to inoculate several mice, but none of the injected animals develop
rhinosporidiosis. Later, Thankamani and Lipin-Dev98 investigated
also the viability of R. seeberi and its developmental stages in
a 10 year-old refrigerated sample. These authors claimed that
they have isolated again the same organism (UMH.48) described
earlier by Thankamani.99 They also illustrated with figures the findings encountered in this archival sample confirming Thankamani99
findings.
To validate the morphological data Thankamani and LipinDev97 recently conducted molecular analysis using genomic DNA
extracted from the original UMH.48 strain (kept on agar slants for 2
years) and directly from new biopsied tissues containing R. seeberi
sporangia. However, the authors did not mention how many clinical samples were processed. They amplified the complete internal
transcriber spacers (ITS) rDNA using the universal primers ITS5 and
ITS4 in both the UMH.48 and the clinical samples.93 The authors
were pleased to find out that the DNA sequence of the strain
UMH.48 and that obtained from biopsied tissue sample with rhinosporidiosis shared 100% identity to each other. They went on
to say that the “. . .99% identity between our isolate UMH.48 and
the fungal DNA from rhinosporidiosis biopsy with respect to the
18S rDNA gene sequence categorically reaffirms that UMH.48 is
the etiology of human rhinosporidiosis.”. Based on these data, they
concluded that the strain UMH.48 seems to be a dimorphic fungus
resembling Synchytrium species with the morphological features
of R. seeberi as described by Seeber88 and Ashworth.16 When the
sequences of both samples (UMH.48 and the biopsied tissue sample) were used to interrogate the database using BLAST analysis,
99% identity was found with the ascomycete Colletotrichum truncatum (Glomerella teleomorphic stage).97 The authors agreed that
this finding was in direct contradiction with the morphological features reported on the original UMH.48 strain.99 But they did not
elaborate more. Instead, they compared their sequences (accession numbers JN807465 and JN807466) with chytridiomycetes and
mesomycetozoans sequences. As expected from the BLAST analysis, the sequences used by Thankamani and Lipin-Dev97 showed
little identity in common with the members of these two groups
of microbes. Although the authors did not conduct phylogenetic
analysis with their DNA sequences, they stated that their study
confirmed R. seeberi as a fungus.
Critical review of R. seeberi’s current interpretations
Is R. seeberi a cyanobacterium? The cyanobacteria comprise unicellular prokaryotic microbes currently grouped in five orders:
Chroococcales, Pleurocapsales, Oscillatoriales, Nostocales, and
Stigonematales.56,57 Members of the Chroococcales possess single
191
Fig. 5. Panel A shows EM photographs of Microcystis’ nanocytes with gas vesicles (gv) (arrows) (gv-l = long, gv-c = circular) a distinctive feature of the genus, whereas Panel B
depicts a nanocyte without gas vesicles (Bars = 500 nm) (Courtesy of Dr. N. Tandeau de Marsac. Used with permission of J Bacteriol). The nanocytes with three peptidoglycan
layers are observed in both panels. The lower section of Fig. 4 shows a cartoon demonstrating the typical binary fission division in more than one plane, typical of the genus
Microcystis (a–c). The formation of large amorphous colonies by the aggregation of nearby cocci (d and e) is a common characteristic of the genus. The picture labeled “e”
shows an actually colony of M. aeruginosa (Courtesy of Mark Schneegurt) (Bar = 50 (␮m). Note an almost invisible mucilage matrix holding the nanocytes. Disintegration of
old colonies (f) is necessary for the colonization of new environmental sites.
spherical cocci (nanocytes) with homogeneous blue-green, grayish,
or yellowish content and reproduce by binary fission in one, two,
or three planes at right angles or in irregular planes (Fig. 5a–c, this
review). Cells (cocci) can reach 0.5–30 ␮m in diameter and form
aggregate of nanocytes (colonies >200 ␮m in diameter) displaying
semispherical, discoid, or irregular shape formations depending on
the planes of division (Fig. 5d and e, this review).80 The aggregated
cocci (colonies) lack the presence of a true cell wall, but the colony
secrets an extracellular slime that holds the nanocytes together.
The reproduction of new colonies occurs by disintegration of the
old colonies into small cluster of cells (Fig. 5f, this review) and new
colonies are formed by the active aggregation of cocci (Fig. 5e, this
review).57
Members in the genus Microcystis, including M. aeruginosa,
are classified within the Chroococcales.74,109 M. aeruginosa is a
common cyanobacterium found worldwide in fresh water environments. In nature M. aeruginosa cell division occurs by binary fission
in three perpendicular planes and in a regular cubic arrangement
of cocci (nanocytes), in the manner of those depicted in Fig. 5, this
review.56,57,74,80 The nanocytes, by binary fission, develop into their
final size (3–15 ␮m) and shape (spherical) before the next cell division, but lack individual mucilage sheaths. Blue-green colonies are
formed by the aggregation of several nanocytes forming spherical,
oval, lobate or irregular colonies held together by a mucilage matrix
(Fig. 5d and e, this review).57,79 Multiplication is by disintegration
of the old colonies into cluster of nanocytes or single dispersed
coccus.57,62,79,109
The cell developmental features of the genus Microcystis in
nature are in contrast with the parasitic features of R. seeberi
in infected hosts. For instance, in the infected host R. seeberi mature
spherical cells, cleave their nuclei to develop numerous endoconidia (see above).37,39,45–47,73,100 R. seeberi’s endoconidia (5–10 ␮m)
are then released through a pore, and once in the infected tissues each endoconidium increases hundreds of times its original
size (>450 ␮m). The enlarged sporangia in turn produces hundreds of new endoconidia that are then released through a pore
and the cycle is repeated (Fig. 4, this review).48,99 By contrast, in
the genus Microcystis: a) nanocyte cell division occurs by binary
192
fission in three perpendicular planes either outside or inside the
colony’s mucilage matrix; b) the shape and size of nanocytes remain
unchanged throughout the entire cell cycle; c) in nature the amorphous or spherical colonies are formed by the aggregation of nearby
nanocytes that became engulfed within a mucilage matrix; and
d) the formation of new amorphous or spherical colonies takes
place only after disintegration of old colonies. So far, there is not
a single report describing: a) R. seeberi producing endoconidia by
perpendicular binary fission in three planes; b) unchanging sizes
of released R. seeberi endoconidia; c) formation of R. seeberi mature
sporangia by the aggregation of nearby endoconidia; d) formation
of an exogenous mucilage matrix in mature sporangia, or d) the
disintegration of R. seeberi mature sporangia to form new mature
spherical cells by aggregation. None of these Microcystis cell developmental features have been ever encountered in the tissues of the
infected patients with rhinosporidiosis.2,16,46,47,54,59,63,75,87,101
According to Alhuwalia6 the morphological features of the
genus Microcystis in nature and R. seeberi in the host’s infected
tissues are almost identical. However, a close inspection of the
Microcystis nanocyte and the endoconidium of R. seeberi revealed
contrasting differences. In addition to the presence of a distinctive
nucleus in R. seeberi endoconidia as reported by many,2,16,40,54,59,100
Sleyter et al.,92 reported that Microcystis nanocytes in EM have
three uniform peptidoglycan layers and an outer proteinaceous
S-layer. In contrast, the endoconidia of R. seeberi lack these cell
wall features (see above) and at this stage undergo dramatic
changes to the thickness of its bilaminated cell wall, a feature
absent in the genus Microcystis.62,92 Furthermore, the most distinctive feature of Microcystis’ nanocytes using EM is the presence
of gas vesicles (GV) within the nanocytes (Fig. 5A, this review).44
This feature is considered almost diagnostic for the genus Microcystis, in which these structures play a key role during its life
cycle in nature. In EM preparations GV are long conical or circular (depending on the angle of the section) hollow gas-filled
structures44,74,101 (Fig. 5A and B, this review), which appear
similar to empty bee-like honey combs in several places in a
nanocyte. However, and in spite of having such unique identifying characteristics, no studies have documented the presence of
entities similar to GV in R. seeberi endoconidia in patients with
rhinosporidiosis.2,13,16,45–48,54,59,87,95–101 Although it could still be
argued that the GV are only needed in nature and thus are not
be displayed in other environments. Arguing against this particular possibility are the numerous other ultrastructural features of
nanocytes devoid of GV (Fig. 5B, this review) that are equally different from those associated with R. seeberi endoconidia (Fig. 2A,
this review).44,74,104 In summary, the reviewed literature indicates that R. seeberi’s endoconidia do not divide into three planes
and lack GV or GV-like structures.2,16,46,59,75,87,100,101 Although
the intracellular vesicles in nanocytes, which are believed to be
pools of reserve substances, have been also found in R. seeberi,
these structures also possess key differences as well (Fig. 5A and
B).45,47,62,101,104
The microscopic arrangement of the nanocytes in colonies
of Microcystis is distinctively different when compared with the
arrangement of endoconidia found in R. seeberi. Also, Microcystis
colonies do not have a true cell wall and the nanocytes are held
together by an amorphous mucilage matrix; whereas R. seeberi has
a well developed thin cell wall, which contains three inner layers holding the endoconidia (Fig. 1, this review).2,16,42,54,59,71 In
addition, R. seeberi mature sporangia with endoconidia are always
characterized by their spherical shapes, whereas the aggregated
colonies of the genus Microcystis display multiple morphologies
depending on the environmental conditions.57,62,79,109 Together
with the presence of a nucleus, these contrasting cell developmental differences distinctly differentiate Microcystis species from the
genus Rhinosporidium.
Does R. seeberi possess rDNA from a cyanobacterium ancestor? The
original proposal of Alhuwalia9 was based on the morphological
attributes that Microcystis cells shared, according to this author,
with R. seeberi. However, they did not perform EM studies to confirm their theory. Instead, a molecular study to prove that DNA
of Microcystis sp. was present in the tissue of humans with rhinosporidiosis was carried out.31 This preliminary molecular study
concluded that the DNA band pattern found in the water containing
Microcystis cells matched the patterns observed from clinical samples and thus was used as proof of the “existence of a pathogenic
strain of Microcystis for humans”.31 A drawback of this preliminary study was the lack of controls. Two years later Dhaulakhamdi
et al.,30 published yet another molecular study. This time genomic
DNA samples from: a) pure culture of M. aeruginosa, b) genomic DNA from putatively pure R. seeberi sporangia incubated in cell
culture, and c) sporangia collected directly from clinical samples
(at least 12 different clinical samples) were subjected to PCR-based
analysis. They concluded that the amplified sequence has the signature for the 16S rDNA gene of cyanobacteria and thus, confirmed
the prokaryotic nature of R. seeberi. However, when the amplified
prokaryotic DNA from tissue samples infected with R. seeberi was
compared to the homologous DNA sequence of M. aeruginosa only
moderate identity was found. Sadly, the authors did not make further comments about this particular finding. Moreover, when the
16S rDNA deposited in the database was used in BLAST analysis
the authors found 98% of identity with plastids present in flowering plants. This fact prompted Alhuwalia et al.,4 to justify that
the relationship between R. seeberi 16S rDNA and those present in
plants explain R. seeberi’s typical auto fluorescence at 510–540 nm
wavelengths, a characteristic of photosynthetic microbes.
Based on ultra-structural studies depicting the presence of a true
nucleus (see above) and the photographic evidence presented in
this review, certainly R. seeberi is a eukaryotic microbe. If R. seeberi,
in addition to being a eukaryotic microbe, possesses prokaryotic
rDNA, then this finding is intriguing. Assuming that the original
PCR experiments were free of environmental contaminants,30 this
finding may be interpreted as the acquisition of prokaryote plastids
by endosymbiosis; but does not necessarily means that R. seeberi
is a cyanobacterium. On the contrary, this finding could provide
new insights into the R. seeberi evolutionary history leading to the
acquisition of such plastids from a cyanobacterium ancestor. Thus,
endosymbiotic acquisition of a plastid in R. seeberi could be a possible explanation to this discrepancy. This scenario is plausible since
the transfer of such plastids from ancient cyanobacteria ancestors
to eukaryotic microbes have been well documented.28,33,36,56,86
So far, plastids transferred from cyanobacteria to eukaryotic cells
have been found in algae,33 apicomplexans,51 cryptomonadas,29
dinoflagellates,36 and plants.28 If R. seeberi had acquired such plastids, it will be the first protist closely related to fungi and animals
with this particular feature.
Is R. seeberi a eukaryote Mesomycetozoa linked to aquatic fish
parasites? Although the presence of a nucleus is always a distinctive
feature of eukaryotic organisms, the finding of this organelle to
support a R. seeberi taxonomic link to the eukaryotes has proved
to be difficult (see below) for various reasons: 1) because R. seeberi
resists culture most studies have been conducted in the tissues of its
infected host. Thus, depending on the tissue sectioning plane, the
finding of a nucleus has been always fortuitous, 2) few investigators
are familiar with the microscopic and ultra-structural characteristics of R. seeberi’s nuclear division during its life cycle (see above),
3) the morphology of R. seeberi’s nucleus and nucleolus in microscopic and EM preparations has yet to be properly defined, and
4) the presence of nuclei, mitochondria, nuclear membrane
and other entities within the spherical structures of R. seeberi are
directly affected by the protocols followed during formalin fixation
treatment. Thus, investigators at different times have reached
varied positions and applied different interpretations to their
observations.2,9,16,47,53,54,59,64,75,87,101 However, with the advent of
molecular methodologies a new door was opened. Good examples
of the application of this new technology to the phylogeny of
R. seeberi are the studies of Frederick et al.,35 Herr et al.,42 Pereira
et al.,82 Silva et al.,91 and Suh et al.94 These independent teams
all have reported that they have molecular data indicating that
the studied epitopes of R. seeberi genomic DNA extracted from
humans and animals, place this unique pathogen with previously
unclassified protistan fish parasites. Perhaps the most important
aspect of the finding was that two independent research groups,
utilizing two different mammalian species with rhinosporidiosis,
both amplified an 18S rDNA molecule sharing 100% identity,
and both groups obtained identical phylogenetic trees.35,42 This
finding leads both groups to believe that only one species, R. seeberi,
was affecting the various animals having rhinosporidiosis.68
Alhuwalia4 stated that a strong argument against the eukaryotic
theory of R. seeberi was that Herr et al.42 and Fredricks et al.,35 probably PCR-amplified DNA from endoconidia of the Dermocystidium
spp. commonly found in aquatic environments.3,68 She predicted
that the amplification of Dermocystidium 18S SSU rDNA was
related to a few cells present in the tissues of humans and animals
previously in contact with ponds containing putative fish infected
with Dermocystidium spp. However, Fredricks et al.,35 clearly
show that the specific Rhino-fw and Rhino-rev primers did not
amplify D. salmonis genomic DNA used as a control, a datum that
argues against this claim. Moreover, when the 18S SSU rDNA from
D. salmonis spp. was used in BLAST analysis by us, key DNA
differences were detected among these two groups of pathogens
(Fig. 6, this review). Thus, the argument that the human tissue
193
used as negative controls by Herr et al.,42 and Fredricks et al.,35
yielded negative PCR results, because the selected individuals had
no previous exposure to environments containing spores of Dermocystidium, is untenable. Another finding that validates Fredricks
et al.35 conclusions, is that their DNA in situ probe detected
R. seeberi DNA inside the sporangia in histological preparations
(blue color), but not in the negative controls (including D. salmonis),
ruling out autofluorescence.90
The EM Fig. 2 of Herr et al.,42 and Fig. 7 this review showed
an IS with one or several nuclei and some mitochondria. Those
supporting the prokaryotic nature of R. seeberi6,9 had argued that
the nuclei and mitochondria depicted by Herr et al.,42 and that
reported by others2,16,42,45,46,59,87 were more likely of human origin or “encompassing naked prokaryotic DNA”. These positions are
understandable because of the common finding of inflammatory
cells residing inside the cell wall of dead sporangia, such as the
one depicted in Fig. 8 of this review. However, the description of
the R. seeberi’s nuclear features in Figs. 2 and 3 (this review) show
substantial differences with the nuclei found in mammalian cells.
For instance, Acevedo,2 Ashworth,16 and Kurunaratne59 described
R. seeberi in H&E-stained preparations having a reddish spherical
nucleus with a typical nucleolus surrounded by a delicate nuclear
membrane similar to those depicted in Figs. 2 and 3 (this review).
In multinucleated IS of R. seeberi (<150 ␮m), nuclei with prominent
nucleoli were identical to those found in single endoconidia and
JS59 (Fig. 2A, this review). Alhuwalia5 arguments against the
identity of the nucleus shown in the study of Herr et al.,42 were
that these investigators confused a nucleus from the host tissue
as R. seeberi’s own nucleus. A close inspection of the host nuclei
in histopathology (H&E) and EM (Figs. 2, 3 and 8, this review),
A
Dermocystidium sp.
Dermocystidium sp.
Dermocystidium salmonis
Amphibiocystidium ranae
Sphaerothecum destruens
Dermocystidium percae
AF158369
AY372365
AF118851
AF399715
DSU21336
AF533950
DSU21337
AY550245
AY692319
FN996445
AY267345
AF533944
AF533949
142-TAAAAAACCAATGCGG-ATCTTTTGGGTCCGGTTCTTT-179
163
163-TAAAAAACCAAT
TAAAAAACCAATGCGG
GCGG-ATCTTTT
ATCTTTTGGGTCCGGTTCTTT
TT-200
200
215-TAAAAAACCAATGCGGGCTCTTGT-GGTCCGGTTCTTT-252
213-TAAAAAACCAATGCGGGC----CTCGGTCCGGTTGTTT-240
201-TAAAAAACCAATGCGGGC----TTCGGTCCGGTTGTTT-238
169-TAAAAAACCAATGCGA----ATTTCGGTTCGGTTCTTT-206
169-TAAAAAACCAATGCGA----ATTTCGGTTCGGTTCTTT-206
196-TAAAAAACCAATGCGA---GTTAACCCTC-GGTTTCTT-233
165-TAAAAAACCAATGCGA---GTTAACCCTC-GGTTTCTT-195
195-TAAAAAACCAATGCGG-ATCTTTTGGGTCCGGTT-CTT-225
195-TAAAAAACCAATGCGG-ATCTTTTGGGTCCGGTT-CTT-225
B
Rhi
idi
seeber
b i
hinosporidium
Dermocystidium sp.
Dermocystidium sp.
Dermocystidium salmonis
AF158369
AY372365
AF118851
AF399715
DSU21336
AF533950
DSU21337
AY550245
AY692319
FN996445
AY267345
AF533944
AF533949
Identity
100%
100%
100%
99%
98%
98%
98%
98%
98%
95%
95%
92%
92%
Differences
0
0
0
3
23
29
35
37
37
87
85
90
92
Gaps
0
0
0
2
7
6
10
10
11
27
29
63
63
Fig. 6. The partial 18S SSU rDNA sequences of several members in the Dermocystida are shown in Panel A. The differences that R. seeberi displayed at the DNA level when
compared to Dermocystidium spp. and to the other members of the group are depicted. The DNA sequences accession numbers are placed after the species name. The numbers
at the right and left section of each DNA sequences represent the location at molecular level of the selected epitope available at the NCBI database (Panel A). Panel B shows
the percentage of identity, the numbers nucleotide differences, and the number of gaps encountered in the 18S SSU rDNA molecules of the 13 DNA sequences in panel A. The
figure illustrates the differences encountered at DNA level between R. seeberi and the other members of the Dermocystida to support the concept7,8 that this mammalian
pathogen can be differentiated using DNA sequence patterns from the genus Dermocystidum.
194
Fig. 7. The electromicrograph in panel A shows an enlargement of a juvenile sporangium with a typical nucleus and a prominent nucleolus (Bar = 8.0 (␮m). Note the
nuclear membrane surrounding R. seeberi’s chromatin. Details of the nuclear membrane and the nucleolus can be observed in panel B (Bar = 2.0 (␮m). In this EM close
up the presence of several vesicles is also observed around the nucleus. Panels A and
B and Figs. 2 and 3 (this review) give details on the typical morphological features
of R. seeberi nuclei and establish without doubt that R. seeberi is a true eukaryote
microbe.
however reveals that the host nuclei are strongly basophilic and
their chromatin is condensed around the edges (Fig. 8, this review),
whereas R. seeberi nuclei are smaller, pale, slightly eosinophilic
and have a prominent nucleolus located at the center or slightly
toward their edges (Figs. 2 and 3, this review).
In Fig. 2B of this review the presence of three JS are depicted.
This figure shows that the finding of a nucleus greatly depends
on the plane of the sectioning. The JS on the left lacks a nucleus,
whereas the other two sporangia show the presence of nuclei. The
two JS on the right section of the figure depict the sectioning of
the nucleus at different plains. Thus the nucleus present on the
JS at the center appears smaller than it really is compared to the
one that is closer to the right edge. More importantly, the presence
of a macrophage nucleus at the top of the figure (arrow) clearly
illustrates the contrasting differences between R. seeberi nuclei and
in that of the infected hosts. Interestingly, the description of early
investigators such as Acevedo,2 Kurunaratne,59 Kennedy et al.,54
and Thianprasit and Thangerngpol,100 all agree with the nuclear
morphological features depicted by Herr et al.,42 in their Fig. 2,
and thus supporting the interpretations of the nuclei described
in this review (Figs. 2A, B, 3A, B, 7A, B, and 8A, B). We contend
that Alhuwalia’s5 argument about the presence of “encompassing
naked prokaryotic DNA” in the pictures provided by Herr et al.,42
are unsubstantiated. Moreover, the chances that a circular prokaryote chromosome in an EM preparation could be sectioned in a way
that shows the whole molecule in one section would be extremely
difficult. In fact, it is more conceivable that the figures of Herr
et al.,42 depicting several nuclei of different sizes surrounded by
a nuclear membrane, comprise spherical structures sectioned at
different points rather than a circular pattern entirely sectioned in
one plane.
Is R. seeberi a pathogen that belong to the kingdom Fungi? The
order Chytridiales (Chytridiomycetes) are divided into seven families: Olpidiaceae, Synchytriaceae, Achlygetonaceae, Rhizideaceae,
Entophlyctaceae, Cladochytriaceae and Physodermataceae.49
Primitive species in this order are mostly holocarpic, whereas
advanced species tend to be eucarpic, forming spherical cells.49 In
the latter group, a system of rhizoids is present to anchor the cell to
its host. The Chytridiales are intracellular pathogens with asexual
reproduction by the development of uniflagellated planospores
formed inside sporangia with one or two pore (papillae).49,51
Some species developed a typical cap at the tip of the pore termed
operculum, but most species lack these opercula.49 In the latter
case the flagellate spores are release through a pore in the cell wall
or by the formation of an exit canal.49,52 In most species the sexual
stage is yet to be found.49
The family Synchytriaceae, where Synchytrium species are
located, comprises intracellular parasites forming uniflagellate
planospores. They are holocarpic Chytridiales lacking opercula, a
feature in common with the family Olpidiaceae.49–52 When
a Synchytrium uniflagellated planospores reaches the infected
Fig. 8. Panels A (Bar = 10 (␮m) and B (Bar = 10 (␮m) show two different histological sections stained with H&E depicting dead sporangia of R. seeberi invaded by the host’s
inflammatory cells. The dead sporangia in panels A and B are commonly found in histological preparations. The nuclei observed inside the dead sporangia are indistinguishable
from the nuclei of the host’s inflammatory cells found around these sporangia (Panels A and B) and different to the R. seeberi’s nuclei depicted in Fig. 3. The figure shows that
R. seeberi nuclear morphological features can be properly differentiated from that of the host’s own nuclei.
hosts an amoeboid spore is formed and it sinks to the bottom
of the parasitized cell where it germinates forming a structure
termed prosorus.51,52 By nuclear divisions the prosorus is emptied and once outside, the cytoplasmic mass becomes the sorus
with several multinucleated cells that at maturity transform into a
sporangium with numerous planospores. It usually remains
attached to the empty structure where the prosorus used to be
located. If water is present, motile planospores are formed and
released. The planospores have only one flagellum and exit the
sporangium through a pore searching for a new host. Once they
localize the host, the zoospores attach to the cell dissolving a section
of the host’s cell wall and penetrate leaving the flagellum outside
the host.51 If water is scarce, the spores transform in planogametes
and fuse in pairs to form zygotes.51,52 The genus Synchytrium has
been divided by Karling52 in several subgenera, but they will not
be discussed in this review.
The isolation of fungi from cases of rhinosporidiosis is not
new. It has been recorded several times.14,58,59,69 The report of
Thankamani99 is peculiar because the report mentioned the isolation of the same fungal organism from 15 different patients and
from a ten-year old frozen sample.99 They described the isolation
of a spherical fungus with all the morphological features that the
author believes are very similar to that in the Chytridiales. The 24
figures describing the fungus in this report are not clear enough to
confirm the author’s claim. However, this is not the major problem
of the report. When we compared the above discussed morphological features of R. seeberi with the description of Thankamani99
several inconsistencies were evident. First, there is not a single
rhinosporidiosis report describing the formation prosorus-like
structures emerging to form multinucleated sorus leaving behind
empty structures (resting prosorus), to which it remains attached
(as in the Chytridiales). Second, R. seeberi mature sporangia release
endoconidia through a pore that do not develop flagella in the presence of water.16,71 In contrast, all chytridiomycetes in the presence
of water release planospores through a pore with a single flagellum
that is used to swim and find a new host. And third, Chytridiomycetes are mostly intracellular parasites, whereas R. seeberi cells
cannot be intracellular in mammalian hosts by its enormous size.
Moreover, the ultrastructural features of the cell wall in R. seeberi
parasitic stages are different to that described in the members of the
Chytridiales.53–57 And finally, the report of Acevedo,2 Ashworth16
and Kurunaratne59 of synchronized multiple nuclear divisions and
the presence of mitotic figures in R. seeberi clinical samples are in
direct contrast with the nuclear division and life cycle displayed by
the Chrytridiomycetes and ascomycetes in their parasitic stages.53
Although Thankamani and Lipin-Dev97 stated that they have
sequenced the 18S SSU rDNA of the UHM.48 strain, they amplified and worked with the ITS rDNA sequences. After BLAST analysis
using the amplified DNA sequences of the original strain (UMH.48)
and that recovered from a clinical sample, the authors were surprised to find out that their isolate was not a Chytridiomycete but
an ascomycete in the genus Colletotrichum. This could well suggest that the morphological description of the original isolate was
not accurate, or that the original strain contaminated later with
Colletotrichum spores. Interestingly, the clinical sample amplified
the same ITS DNA sequence as that in the UMH.48 strain. This was
interpreted by the authors as the confirmation that this fungus is
the etiologic agent of rhinosporidiosis. However, they did not discuss the possibility of cross-contamination between DNA samples,
which could also explain the result.
In the discussion section the authors mentioned that those
supporting the mesomycetozoa theory did not use the scientific
method, but their own opinions on the subject to classify R. seeberi in the mesomycetozoa. However, the data generated by those
teams are supported by multiple controlled phylogenetic analyses from numerous collected samples in humans and animals with
195
rhinosporidiosis.35,42,87 The NCBI database contains many DNA
sequences recovered from humans in India, Sri Lanka, the USA and
Venezuela, as well as DNA sequences from several swans, dogs
and cats with rhinosporidiosis. These sequences link R. seeberi with
strong phylogenetic support to the mesomycetozoans. Therefore,
the hypothesis that R. seeberi is a mesomycetozoa is based, no on
a single case, but on scientific data collected by several independent teams from numerous mammalian and bird hosts with proven
rhinosporidiosis.35,42,87,94
One of the main problems of those supporting the fungal theory of R. seeberi is the lack of controls. As per Mendoza et al.,69
the isolation of genomic DNA from uncultivated pathogens is
difficult. Cross contamination with normal flora and environmental fungal and bacteria microbes is common.69 This had directed
some investigators to misleading views about the taxonomy of
well known uncultivated microbes.65 One of the controls missed
by Thankamani and Lipin-Dev97 is the use of Fredricks et al.,35
specific primers for R. seeberi. This control would allow the
authors to verify if the clinical sample under investigation possesses the same sequence as those reported in Indian human
cases of rhinosporidiosis.91 This control would also suggest to
the authors that the isolated fungal strain was or not a contaminant.
Two other topics mentioned by Thankamani99 deserve special
attention. The UMH.48 strain was isolated on media commonly
used in the laboratory: blood agar nutrient agar and SDA. The samples were incubated at room temperature, because the UMH.48
strain did not grow at 37 ◦ C. This strain also failed to infect
mice. There are no reports describing a mammalian pathogen
microbe that grows at room temperature but resists 37 ◦ C. Thus,
based on this feature it could be concluded that the original
strain was not a pathogenic fungus, but a common contaminant, probably Colletotrichum sp. Also, contrary to Thankamani
and Lipin-Dev97 belief, Colletotrichum spp. are not dimorphic
fungi as those studied in medical mycology. The authors supporting the fungal theory did not ask: why the early investigators
failed to recover R. seeberi in culture? And why we succeed
in 100% of the clinical samples? There seems to be a major
inconsistency with the sudden success in recovering R. seeberi
in culture. Thankamani and Lipin-Dev97 and Thankamani99 did
not use new culture media or novel methodologies that have
not been previously used by others. So, why other investigators had failed to culture R. seeberi using an identical approach?
The answer seems to point more likely to an environmental contamination. So far all fungal isolates recovered from
cases of rhinosporidiosis proved to be environmental fungal
contaminants, and their apparent similarities shared with R. seeberi the lack of taxonomic expertise of those supporting these
claims.14
The morphological and phylogenetic attributes of R. seeberi are in agreement with that displayed by the members
of the Mesomycetozoa. The Class Mesomycetozoa, recently
revised,68 comprises two Orders: Dermocystida and Icthyophonida. According to Herr et al.,42 and Fredricks et al.,35 R. seeberi
is located within the Dermocystida along with Amphibiocystidium,
Amphibiothecum, Dermocystidium, and Sphaerothecum.3,68 The
group is characterized by the presence of spherical unicellular
microbes with the capacity to infect fishes, amphibians, birds and
mammals.68 The main attributes of the group includes: 1) their
typical spherical structures containing hundreds of endoconidia,
2) their long history of inclusion in different groups of microbes,
including the fungi, 3) the resistance of some to culture, 4) all being
aquatic microbes, and 5) all being pathogens of animals. But, the
most striking similarity among them resides in their DNA make
up. Phylogenetic analyses have shown that members of this group
share similar evolutionary paths and together with members of
196
the Ichthyophonida have a common ancestor located at the point
where animals diverged from the fungal boundary.35,42,68,82
The finding that Ichthyophonus hoferi and R. seeberi both posses
chitin synthase genes,41,93 as do other protistal microbes,76 was
used to emend the term Choanozoa to Mesomycetozoa and also
revealed another contrasting evolutionary feature that is not
present in prokaryotic cells.3,56,68 In this context, phylogenetic
analyses have shown (Fig. 9, this review) that R. seeberi forms
a well supported sister taxon with A. ranae and with Dermocystidium species.82 This group in turn is sister to S. destruens and
Dermocystidium percae. Pereira et al.,82 suggested that D. percae was
probably not a Dermocystidium species, but a member of an entirely
new genus, and proposed the name Dermothecum. Although the
members in the Dermocystida shared similar DNA sequences, they
nonetheless posses distinctive differences that are easily demonstrated using CLUSTAL and phylogenetic analysis as those shown in
Figs. 6 and 9 of this review. This unequivocally demonstrates that
the species of Dermocystidum and the other members of the Dermocystida can be properly separated using sequences alignment,
phylogenetic analyses, and DNA probes. Taken together, these facts
all further contradict Alhuwalia4,5 claims that Herr et al.,42 and
Fredricks et al.,35 amplified DNA from samples putatively contaminated with Dermocystidium spp.
Mendoza et al.,68 and Pereira et al.,82 revised the microscopic and ultrastructural attributes of some members in the
Dermocystida. The presence of nuclei very similar to those
described by Acevedo,2 Kurunaratne,59 Savino and Margo,87 Thianprasit and Thangerngpol100 in humans, and Kennedy et al.,54 in
swans infected by R. seeberi was also recorded by Pereira et al.,82 in
frogs harboring A. ranae. Pereira et al.,82 showed a number of striking microscopic and molecular similarities that A. ranae shares with
R. seeberi. Both pathogens not only develop spherules with numerous endoconidia, but both have identical ultrastructural features in
common. For example, in Fig. 3A and B Pereira et al.,82 showed the
morphological features of these pathogens in EM. They call attention to the fact that R. seeberi and A. ranae develop identical electron
dense bodies (EDBs), and similar vesicles within the endoconidia.
They also mention that the cell wall of the endoconidia and the
mature sporangia of A. ranae share a number of features in common with R. seeberi. In panel B of Fig. 3 they show in both pathogens
nuclei with a prominent nucleolus inside a mature endoconidium
containing EDBs. They also stated that the presence of a nucleus in
A. ranae endoconidia is very difficult to document, another interesting attribute in common with R. seeberi. In addition, S. destruens and
Amphibiothecum penneri both develop spherical structures with
endoconidia, and their ultrastructural features also share striking
similarities with R. seeberi, but not with the Chytridiomycetes. Thus
it is not surprising that Laveran and Petit63 and later Carini22 called
the attention on the similarities of Dermosporidium from frogs (currently Amphibiocystidium) and R. seeberi.
In 2005 Silva et al.,91 studying the ITS DNA sequences of R. seeberi extracted from a dog in the USA, several humans in Sri Lanka
and India, and at least two swans in Florida, reported that the
phylogenetic analysis using these gene sequences suggested
the possibility of several species specific strains within R. seeberi.
The ITS DNA sequences of R. seeberi in this study again reinforce the concept that R. seeberi is indeed a eukaryotic pathogen
with capabilities to infect mammals and birds. These phylogenetic analysis using the ITS sequences derived from the DNA of
R. seeberi extracted from humans, swans, and dogs showed that
they clustered in three strongly supported taxons, thus indicating the presence of at least three possible species-specific
strains.
The discussed phylogenetic analyses and the revised microscopic and the EM morphological features of members in the
Dermocystida are in agreement with that described in the parasitic
life cycle of R. seeberi. Sadly, those involved with the prokaryotic
and fungal theories did not conduct the appropriate microscopic
and ultra-structural studies on their samples recovered from ponds,
and R. seeberi from infected patients to avoid bias in the interpretation of their data.
R. seeberi is an eukaryote pathogen in the Mesomycetozoa
When Herr et al.,42 Fredricks et al.,35 and more recently
Suh et al.94 found molecular data supporting the view that R.
seeberi was an eukaryote pathogen sharing morphological and
phylogenetic similarities with members of the Dermocystida,
but distant from the fungi, their finding validated 100 years
of similar views.2,16,18,24,45–51,54,59,75,85,87,88,100 More importantly
members of the Mesomycetozoa are eukaryotic aquatic animal
pathogens that manifest as spherical structures producing endoconidia, all features supporting the eukaryotic nature of R. seeberi.64
In contrast, there is not a single cyanobaterium or species of
Chytridiomycetes that so far has been incriminated as causing
disease in animals.21,43,56,57 The observations of a nucleus, as
reported by many,2,16,18,24,38,45–47,54,59,75,85,87,88,94,100 and those
depicted in Figs. 2, 3, and 7 of this review, are the most powerful argument against the hypothesis that M. aeruginosa is the
etiologic agent of rhinosporidiosis. The revised microscopic and
ultra-structural data2,16,18,24,45,53,54,59,75,85,87,88,100,108 and the figures showed in this review have repeatedly shown that Microcystis,
Synchytrium and R. seeberi display fundamental morphological differences in histological preparations and EM studies. Thus, the
claims that R. seeberi is microscopically similar to a cyanobacterium in the genera Microcystis and Synchytrium is scientifically
untenable. In addition, we believe that the PCR amplification of
cyanobateria DNA from tissue samples containing R. seeberi by
Dhaulakhandi et al.,30 does not prove that this unique pathogen
is a prokaryote. On the contrary, it may suggest that R. seeberi
could have acquired plastids sometime during its evolutionary history. Delwiche et al.,28 using DNA sequences and phylogenetic
analysis of the tufA gene have shown that the presence of plastids in the different eukaryote systems has all originated from
cyanobacteria ancestors. Thus, the amplification of cyanobacteria DNA from clinical samples of patients with rhinosporidiosis,30
could imply that R. seeberi had acquired such plastids in the
past.
Those involved in the fungal97–99 and the prokaryote
theories4,6,8,9 make the same mistakes. Both groups isolated
from clinical samples a microbe morphologically similar to
R. seeberi, but failed to ask fundamental questions about the
strength of their finding. The first question they should ask
would be: what is the relationship of the investigated organism
to similar microbes in the tree of life? Once that is determined, the most important issue is if the life cycle traits of
these groups of microbes and their basic morphological features agree with that in R. seeberi. Both groups did not conduct
ultrastructural studies on their organisms; instead they carried
out DNA analysis directly from the isolated microbe and from
clinical samples collected in patients with proven rhinosporidiosis. In both cases they claimed that the same DNA present
in their isolate was also demonstrated in clinical samples and
therefore, the isolated organism was the etiologic agent of rhinosporidiosis. This seems to be a new trend, so we have to be
vigilant to avoid discrepancies and misinterpretation on future
cases.
Finally, we considered of importance to mention that for
the last 18 years those supporting the prokaryotic origin of R.
seeberi have been also adopting multiple views on the nature
of this anomalous pathogen. For instance, Alhuwalia in 19927
15 Rhinosporidium canis AY372365
21 Rhinosporidium cygnus bird AF399715
80 Rhinosporidium seeberi dog AF158369
35
197
R. seeberi
Rhinosporidium seeberi human AF118851
Amphibiocystidium sp newt EF493030
40
82
Amphibiocystidium ranae frog AY550245
40
99 Amphibiocystidium ranae frog AY692319
84 Dermocystidium salmonis fish DSU21337
Dermocystidium sp fish AF533950
96
Dermocystidiumspp.
Dermocystidium sp fish DSU21336
99 Amphibiothecum penneri frog AY772001
Amphibiothecum penneri frog AY772000
Dermocystida sp newt GU232542
78
52
99
91 Dermocystidium parcae fish AF533948
Dermocystidium parcae fish AF533946
99
61
Sphaerothecum destruens fish L29455
Sphaerothecum destruens fish AY267346
99 Sphaerothecum destruens fish AY267345
Sphaerothecum destruens fish AY267344
Sphaeroforma arctica Y16260
99
Ichthyophonus hoferi IHU25637
20
Fig. 9. Parsimony tree aligned 18S SSU rDNA sequences of four R. seeberi and 28 other dermocystidian DNA sequences. The ichthyosporeans Sphaeroforma artica and
Ichthyophonus hoferi were used as outgroup. Numbers above the branches are percentages of 1000 bootstrap-resampled data sets. The scale bar represents substitutions per
nucleotide. The accession numbers are located after the organism’s name. In this phylogenetic tree the four R. seeberi DNA sequences formed a well supported sister taxon
to the five Amphibiocystidium spp. DNA sequences from newts and frogs. The three Dermocystidium species DNA sequences in turn formed a poor supported sister taxon to
Amphibiocystidium and Rhinosporidium. The phylogenetic analysis indicates that Rhinosporidium seeberi shares more phylogenetic features in common with Amphibiocytidium
than to the other members of the Dermocystida. Several strains of Dermocystidium percae were placed in a strongly supported taxon indicating that this pathogen of fish
probably is not a Dermocystidium species. The genus Dermotheca has been suggested by some to differentiate this pathogen of fish from the other Dermocystidium species.
introduced the hypothesis that R. seeberi was “composed of both
plant and human material that is self-assembled in response
to specific function pertaining to its elimination from the tissue”. The same year Alhuwalia8 launched yet another theory.
The author stated that “. . .The so-called sporangium is found to
be a unique body containing residue-loaded lysosomal bodies
(”spores“) for elimination of the system”. She also mentioned that
“Two carbohydrates, namely defective proteoglycans synthesized
intracellularly and an exogenous polysaccharide ingested through
diet of tapioca constitute indigestible material in NB (nodular
bodies) and scw (cellular waste)”. In 1994 Alhuwalia et al.,10
concluded that “. . .Dietary dry tapioca and chronic inflammation in undernourished individuals could lead to rhinosporidiosis”.
Our careful review of these and similar concepts on the origin
of R. seeberi by Alhuwalia and her colleagues suggest lack of
the scientific rigor required to introduce novel concepts in science.
Concluding remarks
We have critically reviewed key microscopic, ultrastructural,
and the molecular studies of the past 110 years and concluded that
R. seeberi: i) does not share the microscopic or ultrastructural features with that found in the genera Microcystis or Synchytrium spp.,
ii) has entirely different life cycle traits than those reported for the
genus Microcystis the chytridiomycetes or the ascomycetous fungi,
iii) possesses a nucleus with a prominent nucleolus and distinctive features distinguishable from those associated with the nuclei
of its infected hosts, iv) is a mesomycetozoa microbe based on
DNA and phylogenetic analysis, v) has microscopic and ultrastructural characteristics that are in agreement with the characteristic
of the Mesomycetozoa, but not with that of the members of lower
fungi or ascomycetes vi) develops mitotic figures in prophase,
metaphase, and anaphase without cytokinesis in its immature sporangia and, thus it is a typical eukaryote microbe taxonomically and
198
phylogenetically different from the fungi, vii) may have acquired
plastid from a cyanobacterium ancestor, and viii) does not develop
uniflagellate cells in any of its developmental stages.
One of the fundamental features discussed at length in this
review is the presence in R. seeberi of a typical nucleus and
the reports of mitotic figures in tissue sections by the early
investigators.2,16,59 The finding of synchronized nuclear division
with the formation of endoconidia only in the latest mature stages,
supports the placement of this unique pathogen in the mesomycetozoa and away from the fungi.2,16,59 We anticipate that the notion
of plastid DNA in R. seeberi will ignite a new interest in the subject,
which could redirect future research efforts in this and other areas
that still need further research.
Conflict of interest
The authors declare no conflict of interest.
Acknowledgements
The authors thank Dr. Paul Szaniszlo for the suggestions and
constructive criticism. The study was supported in part by the
Biomedical Laboratory Diagnostics, Michigan State University.
References
1. Aben-Athar J. Um caso de Rhinosporidiose. Rev Acad Med Belém, Pará Br.
1944;2:3–7.
2. Acevedo P de. Rhinosporidium seeberi: Considerações sôbre morfologia, ciclo
parasitário e posição sistemática. Thesis. Belém, Pará, Brasil: Facultade de
Medicina, Universidade do Pará; 1958. p. 1–108.
3. Adl SM, Simpson AG, Farmer MA, Andersen RA, Anderson OR, Barta JR, et al. The
new higher level classification of eukaryotes with emphasis on the taxonomy
of protists. J Eukaryot Microbiol. 2005;52:399–451.
4. Alhuwalia KB. Sequence analysis of 16S rDNA gene in Rhinosporidium seeberi
shows similarity to plant chloroplast DNA. Bioinformation. 2010;5:89–96.
5. Alhuwalia KB. Causative agent of rhinosporidiosis. Author’s reply. J Clin Microbiol. 2001;39:413–5.
6. Alhuwalia KB. Culture of the organism that causes rhinosporidiosis. J Laryngol
Otol. 1999;113:523–8.
7. Alhuwalia KB. Plant molecule in human disease, a novel association. In:
Wegmann RJ, Wegman MA, editors. Gene regulation and molecular aspects of
muscle, liver, pancreas, connective tissue and plants, vol. 5. Lauven, Belgium:
Peeters Press; 1992. p. 289–92.
8. Alhuwalia KB. New interpretations in rhinosporidiosis, enigmatic disease of
the last nine decades. J Submicrosc Cytol Pathol. 1992;24:109–14.
9. Alhuwalia KB, Maheshwari N, Deka RC. Rhinosporidiosis: a study that resolves
etiologic controversies. Am J Rhinol. 1997;11:479–83.
10. Alhuwalia KB, Sharma N, Kacher SK, Deka RC. Association of tapioca and chronic
inflammation with rhinosporidiosis. Indian J Otolaryngol Head Neck Surg.
1994;3:25–7.
11. Allen FRWK, Dave M. Treatment of rhinosporidiomycosis in man based on sixty
cases. Ind Med Gaz. 1936;71:376–95.
12. Apple DJ. Papillome der conjunktiva bedingt durch rhinosporidiose. Fortschr
Ophthalmol. 1983;79:571–4.
13. Arseculeratne SN. Rhinosporidiosis: what is the cause. Curr Opin Infect Dis.
2005;18:113–8.
14. Arseculeratne SN, Mendoza L. Rhinosporidiosis. In: Merz WG, Hay R, editors.
Topley and Wilson’s microbiology and microbial infections. 10th edition Hodder Arnold: ASM Press; 2005. p. 436–75.
15. Arseculeratne SN, Hussein FN, Atapattu DN, Pathmanathan R. Failure to infect
congenitally immunodeficient SCID and NUDE mice with Rhinosporidium seeberi. Med Mycol. 2000;38:393–5.
16. Ashworth JH. On Rhinosporidium seeberi (Wernicke, 1903) with special reference to its sporulation and affinities. Trans Roy Soc Edin. 1923;53:301–42.
17. Azadeh B, Baghoumian N, El-Bakri OT. Rhinosporidiosis: immunohistochemical and electron microscopic studies. J Laryngol Otol. 1994;108:1048–54.
18. Bader G, Grueber HLE. Histochemical studies on Rhinosporidium seeberi. Virchows Arch Abt A Pathol Anat. 1970;350:76–86.
19. Battie JM. Rhinosporidium kinealyi: a sporozoon of the nasal mucous membrane.
J Pathol Bacteriol. 1906;11:270–5.
20. Belou P. Tratado de Parasitología Animal: Ilustrado con 301 figuras intercaladas
en el texto y 16 láminas en cromo-litografía, vol. 303. Buenos Aires, Argentina:
Arsenio Guidi Buffarini (ed.); 1903. p. 62.
21. Brunberg AK, Bostrom B. Coupling between benthic biomass of Microcystis and
phosphorus release from the sediments of a highly eutrophic lake. Hydrobiologia. 1992;235:375–85.
22. Carini A. Sôbre um parasito smelhante ao Rhinosporidium encontrado em quisto
da pele de uma Hyla. Arq Inst Biol. 1940;11:93–6.
23. Cavalier-Smith T. Neomonada and the origin of animal and fungi. In: Coombs
GH, Vickerman K, Slegh MA, Warren A, editors. Evolutionary Relationships
Among Protozoa, vol. 56. Dordrecht, The Netherlands: Kluwer; 1998. p.
375–407.
24. Ceferri R, Redaelli P, Scatizzi I. Unitá etiologica della malattia di Seeber (Granuloma de Rhinosporidium seeberi) accertata com lo studio di materiali originali.
Boll Soc Med Chir di Pavia. 1936;14:723–45.
25. Cole ST, Eiglmeier K, Parkhill J, James KD, Thomson NR, Wheeler PR, et al.
Massive gene decay in the leprosy bacillus. Nature. 2001;409:1007–11.
26. Cordero EH, Vogelsang EG. El probable modo de infección de las mucosas por
Rhinosporidium. Bol Inst Clin Quir. 1928;28:573–4.
27. Datta S. Rhinosporidium seeberi, its cultivation and identity. J Vet Sci Anim
Husban. 1965;35:1–17.
28. Delwiche CF. Tracing the tread of plastid diversity through the tapestry of life.
Am Nat. 1999;154:S165–77.
29. Delwiche CF, Kuhsel M, Palmer JD. Phylogenetic analysis of tufA sequences
indicates a cyanobacterial origin of all plastids. Mol Phylogenet Evol.
1995;4:110–28.
30. Dhaulakhandi DB, Alhuwalia KB, Ravi AK, Garg LC. Detection of 16S rDNA
gene in round bodies isolated from polyps of rhinosporidiosis. Infect Gen Evol.
2006;6:331–6.
31. Dhaulakhandi DB, Alhuwalia KB, Poduval P, Ravi AK. Comparative RAPD analysis of causative agent of rhinosporidiosis and Microcystis isolated from waters.
Curr Sci. 2004;86:386–8.
32. Dodge CW. Rhinosporidium. In: Fungous diseases of men and other mammals.
St. Louis: The C.V. Mosby Company; 1935. p. 151–152.
33. Douglas SE. Plastid evolution: origins, diversity, trends. Curr Opin Genet Dev.
1998;8:655–61.
34. Fonseca OJM, Lacaz CdaS. Estudo de culturas isoladas de blastomicose queloidiforme (doença de Jorge Lobo). Denominação ao seu agente etiològico. Rev Inst
Med Trop São Paulo. 1971;13:225–51.
35. Fredricks DN, Jolley JA, Lepp PW, Kosek JC, Relman DA. Rhinosporidium seeberi:
a human pathogen from a novel group of aquatic protistan parasites. Emer
Infect Dis. 2000;6:273–82.
36. Gabrielsen TM, Minge MA, Espelund M, Tooming-Klunderud A, Patil V,
Nederbragt AJ, et al. Genome evolution of a tertiary dinoflagellate plastid. PLoS
One. 2011;6:1–8.
37. Gori S, Scasso A. Cytologic and differential diagnosis of rhinosporidiosis. Acta
Cytol. 1994;38:361–6.
38. Grover S. Cytological study of Rhinosporidium seeberi by electron microscopy.
Indian J Pathol Bacteriol. 1974;17:29–32.
39. Grover S. Rhinosporidium seeberi: a preliminary study of the morphology and
life cycle. Sabouraudia. 1970;7:219–31.
40. Herr RA, Mendoza L, Arseculeratne SN, Ajello L. Immunolocalization of an
endogenous antigenic material of Rhinosporidium seeberi expressed only
during mature sporangial development. FEMS Immunol Med Microbiol.
2002;23:2015–212.
41. Herr AR, Ajello L, Mendoza L. Chitin synthase class 2 (CHS2) gene from the
human and animal pathogen Rhinosporidium seeberi. In: Proc. 99th Gen. Meet.
Am. Soc. Microbiol. 1999. p. 296 [Abstr.].
42. Herr RA, Tarcha EJ, Taborda PR, Taylor JW, Ajello L, Mendoza L. Phylogenetic
analysis of Rhinosporidium seeberi’s 18S small-sub unit ribosomal DNA groups
this pathogen among members of the protistan Mesomycetozoa clade. J Clin
Microbiol. 1999;37:2750–4.
43. Janse I, Meima M, Kardinaal EA, Zwart G. High-resolution differentiation of
cyanobacteria by using rRNA-internal transcriber spacer denaturing gradient
gel electrophoresis. Appl Environ Microbiol. 2003;69:6634–43.
44. Jones DD, Jost M. Isolation and chemical characterization of gas vacuole
membranes from Microcystis aeruginosa Kütz. emend. Elenkin Arch Mikrobiol.
1970;70:43–64.
45. Kannan-Kutty M, Teh EC. Rhinosporidium seeberi: an ultrastructural study of its
endosporulation phase and trophocyte phase. Arch Pathol. 1975;99:51–4.
46. Kannan-Kutty M, Teh EC. Rhinosporidium seeberi: an ultrastructural study of its
life cycle. Pathology. 1974;6:63–70.
47. Kannan-Kutty M, Teh EC. Rhinosporidium seeberi: cell wall formation in sporoplasts. Pathology. 1974;6:183–5.
48. Kannan-Kutty M, Gomez JB. The ultrastructure and life history of Rhinosporidium seeberi. S E Asian J Trop Med Publ Health. 1971;2:9–16.
49. Karling JS. Chytridiomycetarum iconographia. An illustrated and brief descriptive guide to the Chytridiomycetes genera with a supplement of the
Hyphochytridiomycetes. New York: J. Cramer; 1977. p. 43–59.
50. Karling JS. Resting spore germination in Synchytrium australe in relation to its
classification. Mycologia. 1955;47:185–92.
51. Karling JS. Prosori in Synchytrium. Bull Torrey Bot Club. 1955;82:218–36.
52. Karling JS. Micromyces and Synchytrium. Mycologia. 1953;45:276–87.
53. Karpova MF. On the morphology of rhinosporidiosis. Mycopathologia.
1964;23:281–6.
54. Kennedy FA, Buggage RR, Ajello L. Rhinosporidiosis: a description of an
unprecedented outbreak of captive swans (Cygnus spp.) and a proposal for
revision of the ontogenic nomenclature of Rhinosporidium seeberi. J Med Vet
Mycol. 1995;33:157–65.
55. Köhler S, Delwiche CF, Denny PW, Tilney LG, Webster P, Wilson RJ, et al.
A plastid of probable green algal origin in apicomplexan parasites. Science.
1997;275:1485–9.
56. Komárek J, Komárková J. Review of the European Microcystis-morphospecies
(Cyanoprokaryotes) from nature. Czech Phycol. 2002;2:1–24.
57. Komárek J, Anagnostidis K. Cyanoprokaryota, 1. Teil: Chroococcales. In: Ettl H,
Gärtner G, Heynig H, Mollenhauer D, editors. Süsswasserflora von Mitteleuropa, Band 19/1. Jena: Gustav Fischer Verlag; 1999. p. 1–548.
58. Krishnamoorthy S, Sreedharan VP, Koshy P, Kumar S, Anilakumari CK,
Krishnamoorty S, et al. Culture of Rhinosporidium seeberi: preliminary report.
J Layngol Otol. 1989;103:178–80.
59. Kurunaratne WAE. Rhinosporidiosis in man. London, Toronto, New York: University of London; 1964. p. 1–146, The Athlone Press.
60. Kurunaratne WA. The pathology of rhinosporidiosis. J Pathol. 1936;1:193–202.
61. Lacaz CdaS. Doença de Jorge Lôbo (Lobo’s Disease). Rev Iberoamer Micol.
1996;13:819–26.
62. Lang NJ. The fine structure of the blue-green algae. Annu Rev Microbiol.
1968;22:15–46.
63. Laveran A, Petit A. Sur une épizootic des truites. C R Acad Sci Paris.
1910;151:421–3.
64. Lengfelder DD, Pospischil A. Rhinosporidiosse beim pferd. Histopathologische
und elecktronenmikroskopische untersuchung eines falles. Zentralbl Veterinaermed [B]. 1980;27:326–39.
65. Levy MG, Meuten DJ, Breitschwerdt EB. Cultivation of Rhinosporidium seeberi
in vitro: interaction with epithelial cells. Science. 1986;234:474–6.
66. Matsuo E. Leprosy as a challenge to science on the ability to decode its enigma.
A hypothesis on how to respond. Jpn J Infect Dis. 2005;58:59–64.
67. Melo MTde. Estudos sôbre o Rhinosporidium seeberi. Thesis. Rio de Janeiro,
Brasil: Estudos Gráficos, Muniz Press; 1946, p. 1–145.
68. Mendoza L, Taylor JW, Ajello L. The class Mesomycetozoea: a heterogeneous
group of microorganisms at the animal-fungal boundary. Annu Rev Microbiol.
2002;56:315–44.
69. Mendoza L, Ajello L, Taylor JW. The taxonomic status of Lacazia loboi and Rhinosporidium seeberi has been finally resolved with the use of molecular tools.
Rev Iberoam Micol. 2001;39:309–14.
70. Mendoza L, Herr RA, Ajello L. Causative agent of Rhinosporidiosis. Author’s
reply. J Clin Microbiol. 2001;39:413–5.
71. Mendoza L, Herr R, Arseculeratne SN, Ajello L. In vitro studies on the mechanisms of endospore release by Rhinosporidium seeberi. Mycopathologia.
1999;148:9–15.
72. Minchin EA, Fantham HB. Rhinosporidium kinealyi n.g., n.sp. a new sporozoan
from the mucous membrane of the septum nasi of man. Q J Microsc Sci.
1905;49:521–32.
73. Mishra RK, Anand K, Kacker SK, Sinhea AN. Ultrastructure of sporangium of
Rhinosporidium seeberi. Ind J Microbiol. 1968;8:215–20.
74. Mlouka A, Comte K, Castets AM, Bouchier C, Tandeau de Marsac N. The gas
vesicle gene cluster from Microcystis aeruginosa and DNA rearrangements that
lead to loss of cell buoyancy. J Bateriol. 2004;186:2355–65.
75. Moses JS, Balachandran C, Singh B, Shanmugan A, Moses D. Rhinosporidium
seeberi: light, phase contrast, fluorescent and scanning electron microscopic
study. Mycopathologia. 1991;114:17–20.
76. Mulisch M. Chitin in protistan organisms. Eur J Prostistol. 1993;29:1–18.
77. O’Kinealy F. Localised psorospermosis of the mucous membrane of the septum
nasi. In: Proceedings of the Laryngology Society of London. 1903. p. 109–12
[Abstr.].
78. Osawa N. Growth of Mycobacterium leprae in vitro. Proc Japan Acad Ser B.
1997;73:144–9.
79. Otsuka S, Suda S, Li RH, Matsumoto S, Watanabe MM. Morphological variability of colonies of Microcystis morphospecies in culture. J Gen Appl Microbiol.
2000;46:39–50.
80. Palinska KA, Krumbein WE. Patterns of growth in coccoid, aggregate forming
cyanobacteria. Ann Bot Fennici. 1998;35:219–27.
81. Pattyn SR. The problem of cultivation of Mycobacterium leprae. A review with
criteria for evaluating recent experiment work. Bull Org Mond Santé, Bull
World Health Org. 1973;49:403–10.
82. Pereira CN, Di Rosa I, Fagotti A, Simoncelli F, Pascolini R, Mendoza L.
The pathogen of frogs and Amphibiocystidium ranae is a member of the
order Dermocystida in the class Mesomycetozoea. J Clin Microbiol. 2005;43:
192–8.
199
83. Prins C, Tange RA, Dingemans KP. Rhinosporidiosis in The Netherlands. A case
report including ultramicroscopic features. ORL J Otorhinolaryngol Relat Spec.
1983;45:237–44.
84. Ragan MA, Goggin CL, Cawthorn RJ, Cerenius L, Jamieson AV, Plourde SM, et al.
A novel clade of protistan parasites near the animal-fungal divergence. Proc
Natl Acad Sci USA. 1996;93:11907–12.
85. Rao SN. Rhinosporidium seeberi. A histochemical study. Indian J Exp Biol.
1966;4:10–4.
86. Sato S. The apicomplexan plastid and its evolution. Cell Mol Life Sci.
2011;68:1285–96.
87. Savino DF, Margo CE. Conjunctival rhinosporidiosis: light and electron microscopic study. Ophthalmology. 1983;90:1482–9.
88. Seeber GR. Un nuevo esporozuario parasito del hombre. Dos casos encontrados
en polypos nasales. Thesis. Buenos Aires, Argentina: Universidad Nacional de
Buenos Aires, Imprenta Libreria Boullosa; 1900.
89. Seeber GR. Rhinosporidium kinealyi et Rhinosporidium seeberi une question de
priorité. Buenos Aires, Argentina: La Ciencia Médica; 1912. p. 10.
90. Senba M, Yamashita H. Autofluorescense of Rhinosporidium seeberi. Am J Clin
Pathol. 1985;83:132.
91. Silva V, Pereira CN, Ajello L, Mendoza L. Molecular evidence for multiple hostspecific strains in the genus Rhinosporidium. J Clin Microbiol. 2005;43:1865–8.
92. Sleyter UB, Messner P, Pum D, Sára M. Crystalline bacterial cell surface layers.
Berlin, Heidelberg. New York, London, Paris, Tokyo: Springer-Verlag; 1988.
93. Spanggaard B, Huss HH, Brescianni J. Morphology of Ichthyophonus
hoferi assessed by light and scanning electron microscopy. J Fish Dis.
1995;18:567–77.
94. Suh LH, Barron J, Dubovy SR, Gaunt ML, Ledee DR, Miller D, et al. Occular rhinosporidiosis presenting as chronic follicular conjunctivitis in a contact lens
wearer. Arch Ophthalmol. 2009;127:1076–7.
95. Teh EC. Surface structure of the endospores of Rhinosporidium seeberi.
Sabouraudia. 1974;12:263–5.
96. Teh EC, Kannan-Kutty M. Rhinosporidium seeberi: spherules and their significance. Pathology. 1975;7:133–7.
97. Thankamani V, Lipin-Dev MS. Rhinosporidium seeberi proven as a fungus for
the first time after a century since its discovery. Res Biotechnol. 2012;3:
41–6.
98. Thankamani V, Lipin-Dev MS. Demonstration of viability and development
stages of 10 years old refrigerated culture of Rhinosporidium seeberi (?) UMH.48
first ever isolated from nasal Rhinosporidiosis yet another milestone, a tribute
to Guillermo Seeber. IJPI’s J Biotech Biother. 2011;1:11–21.
99. Thankamani V. Rhinosporidium seeberi grown in culture. A ray of hope for the
thousands of patients with rhinosporidiosis!!! South Asian J Socio-Political
Studies (SAJOSPS). 2005;5:118–23.
100. Thianprasit M, Thagernpol K. Rhinosporidiosis. Curr Top Med Mycol.
1989;3:64–85.
101. Vanbreuseghem R. Ultrastructure of Rhinosporidium seeberi. Int J Dermatol.
1973;12:20–8.
102. Vilela R, Rosa PS, Belone AF, Taylor JW, Diório SM, Mendoza L. Molecular phylogeny of animal pathogen Lacazia loboi inferred from rDNA and DNA coding
sequences. Mycol Res. 2009;113:851–7.
103. Vilela R, Martins JE, Pereira CN, Melo N, Mendoza L. Molecular study of archival
fungal strains isolated from cases of lacaziosis (Jorge Lobo’s disease). Mycoses.
2007;50:470–4.
104. Walsby AE. Structure and function of gas vacuoles. Bacteriol Rev.
1972;36:1–32.
105. Wernicke R. Ueber einen protozoenbefund bei mycosis fungoindes. Zentralbl
Bakteriol. 1892;12:859–61.
106. Woodard B, Hudson J. Rhinosporidiosis: ultrastructural study on an infection
in South Carolina. South Med J. 1984;77:1587–8.
107. Wright J. A nasal sporozoon (R. kinealyi). New York Med J. 1907;86:1149–53.
108. Xinzhang L, Tinnxi L. Observations on the ultrastructure of Rhinosporidium
seeberi. Chin J Dermatol. 1981;14:72–4.
109. Yamamoto Y, Shiah FK. Relationship between cell growth and frequency of
dividing cells of Microcystis aeruginosa. Plankton Benthos Res. 2010;5:131–5.
110. Zschokke E. Ein Rhinosporidium beim pferd. Schweizer Arch Tierheilkd.
1913;55:641–50.

185-199 - Revista Iberoamericana de Micología

Transcription

Similar documents

Rhinosporidiosis Rhinosporidiosis R. Seeberi (Life Cycle)

EasyPure Genomic DNA Spin Kit

The Masterof

WHY INTERNAL COMMUNICATIONS?

Presentation - Produce Marketing Association

DNA Structure - hrsbstaff.ednet.ns.ca

Cell division

`CSI effect` impacts justice in Tucson

Principios de Biología Molecular

4.2. Tracing populations with Haplogroups